DNA damage-induced gene expression in Saccharomyces ...

R E V I E W A R T I C L E

DNAdamage-inducedgene expression inSaccharomyces cerevisiaeYu Fu, Landon Pastushok & Wei Xiao

Department of Microbiology and Immunology, University of Saskatchewan, Saskatoon, SK, Canada

Correspondence: Wei Xiao, Department of

Microbiology and Immunology, University of

Saskatchewan, 107 Wiggins Road,

Saskatoon, SK, Canada S7N 5E5. Tel.: 11 306

966 4308; fax: 11 306 966 4311;

e-mail: [email protected]

Received 15 February 2008; revised 27 May

2008; accepted 28 May 2008.

First published online 8 July 2008.

DOI:10.1111/j.1574-6976.2008.00126.x

Editor: Martin Kupiec

Keywords

Saccharomyces cerevisiae ; DNA damage;

transcriptional regulation; SOS response;

cell-cycle checkpoint; signal transduction.

Abstract

After exposure to DNA-damaging agents, both prokaryotic and eukaryotic cells

activate stress responses that result in specific alterations in patterns of gene

expression. Bacteria such as Escherichia coli possess both lesion-specific responses

as well as an SOS response to general DNA damage, and the molecular mechanisms

of these responses are well studied. Mechanisms of DNA damage response in lower

eukaryotes such as Saccharomyces cerevisiae are apparently different from those in

bacteria. It becomes clear that many DNA damage-inducible genes are coregulated

by the cell-cycle checkpoint, a signal transduction cascade that coordinates

replication, repair, transcription and cell-cycle progression. On the other hand,

among several well-characterized yeast DNA damage-inducible genes, their

effectors and mechanisms of transcriptional regulation are rather different. This

review attempts to summarize the current state of knowledge on the molecular

mechanisms of DNA damage-induced transcriptional regulation in this model

lower eukaryotic microorganism.

Introduction

DNA is the carrier of genetic information in most organisms.

Any damage to the molecular structure of DNA has the

potential to cause genomic instability, mutagenesis or even cell

death. Unfortunately, DNA damage is unavoidable; DNA is

continually exposed to insults resulting from exogenous and

endogenous DNA-damaging agents as well as challenges posed

by DNA replication. Therefore, it is not surprising that living

organisms have developed numerous pathways to deal with

DNA damage. In response to DNA damage, cells can alter many

of their intracellular processes, including DNA repair, metabo-

lism and cell-cycle progression, for cell survival and mainte-

nance of genomic stability (Friedberg et al., 2006). Such

dynamic responses can be achieved through a variety of

molecular mechanisms such as protein localization, protein

degradation, posttranslational modification and genetic regula-

tion. Generally speaking, altering gene expression is perhaps

one of the most fundamental responses and it is reasonable to

suggest that most, if not all, cellular processes are affected at the

level of transcriptional regulation. In the field of DNA repair

and mutagenesis, the notion that DNA damage causes an

alteration in the expression profile of damage responsive genes

has been an important area of research for many years. It is

expected that the expression profile of a cell following DNA

damage will provide us with critical information on how a cell

protects itself from such stress.

Transcriptional regulation in response to DNA damage has

been studied extensively in model bacteria such as Escherichia

coli. In contrast, such information is relatively scarce in

eukaryotic organisms, from simple lower unicellular eukaryotes

such as Saccharomyces cerevisiae to humans. In this review, we

attempt to summarize recent findings related to gene regulation

in response to DNA damage in lower eukaryotes, particularly

the budding yeast S. cerevisiae, and to compare this response

with the well-studied bacterial SOS response.

Bacterial transcriptional responses to DNAdamage

The SOS response in E. coli

When E. coli cells are subjected to DNA damage, about 48

unlinked genes are coordinately induced through a complex

SOS regulatory network (Courcelle et al., 2001). The in-

creased expression of these SOS regulon genes results in the

elaboration of a set of physiological responses such as an

enhanced capacity for recombination repair and excision

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repair, enhanced mutagenesis (due to error-prone transle-

sion DNA synthesis mediated by PolIV and PolV) and

inhibition of cell division. These responses have been

collectively termed the SOS response (Radman, 1975;

Witkin, 1976; Gottesman, 1981; Little & Mount, 1982).

Current model for transcriptional control of theSOS response

The SOS regulatory network is mainly controlled by two

proteins: RecA and LexA (Radman, 1975; Little & Mount,

1982; Walker, 1984). LexA is a transcriptional repressor that

binds to SOS boxes located near or inside the operator site of

the SOS-induced genes. SOS boxes are often palindromic

structures with a high degree of homology in nucleotide

sequence. An ideal symmetrical consensus sequence 50-

TACTGTATATATATACAGTA-30 was derived from the ana-

lysis of a pool of SOS boxes (Berg, 1988). The sequence

distinction in SOS boxes allows the LexA repressor to bind

to operators with different strengths (Lewis et al., 1994). The

LexA occupancy prevents accessibility to RNA polymerase

so as to inhibit the initiation of transcription. LexA is likely

to interact with DNA via its N-terminal domain, and the

C-terminus of LexA is required for its dimerization. The

dimerization of LexA is essential for its ability to repress

SOS-regulated genes in vivo. Meanwhile, LexA is able to

undergo a slow intramolecular self-cleavage termed auto-

digestion, whose rate significantly increases upon interac-

tion with RecA (Little, 1984, 1991, 1993).

The RecA protein of E. coli has at least three functions

in the SOS response. It not only plays a role in the

transcriptional regulation in response to DNA damage but

also directly participates in translesion synthesis and homo-

logous recombination. Following DNA damage, DNA

synthesis becomes discontinuous and single-strand DNA

(ssDNA) is produced by failed attempts to replicate da-

maged DNA. In the presence of ATP, RecA binds to the

ssDNA region and forms helical RecA-ssDNA nucleopro-

tein filaments. LexA then diffuses to deep grooves in the

RecA-ssDNA filaments and interacts with them in a man-

ner that results in autocatalytic cleavage of LexA at a scissile

peptide bond located between Ala84 and Gly85. Cleavage

of LexA inactivates its ability as a repressor, thus releasing

the SOS regulon genes from transcriptional repression. As

cells begin to recover from the inducing treatment by

various DNA repair and tolerance processes, the regions of

ssDNA disappear, and thus the inducing signal is dimin-

ished. Without the cleavage stimulated by RecA-ssDNA

filaments, the pool of LexA is boosted, which leads to

repression of the transcription of SOS regulon genes and a

return to the uninduced state (Walker, 1984; Friedberg

et al., 2006).

Fine tuning in the induction of the SOS response

The SOS regulatory system provides E. coli with a rapid

transcriptional response to the presence of DNA damage.

Furthermore, during SOS induction, the timing, the dura-

tion and the level of induction are diverse for different LexA-

regulated genes, suggesting a fine-tuning mechanism in the

SOS induction. The fine tuning is possibly determined by

at least four parameters: (1) the binding affinity of LexA

for the SOS box in the operator region; (2) the number of

SOS boxes in the operator region; (3) the location of the

SOS box relative to the promoter; and (4) the strength of

the promoter.

After exposing E. coli cells to DNA-damaging agents,

genes with operators that bind LexA relatively weakly are

the first to turn on fully. For example, uvrA, uvrB, ruvA,

ruvB, recN and sulA are induced within 5 min after 40 J m�2

UV radiation (Courcelle et al., 2001). uvrA and uvrB encode

proteins involved in nucleotide excision repair (NER); recN,

ruvA and ruvB encode proteins used for recombination

repair; while the protein product of the sulA gene can

temporarily arrest cell division to allow bacteria time to

complete the repair of damaged DNA. After the activation of

loosely controlled SOS genes, if the damage cannot be fully

repaired by NER and homologous recombination, genes

with operators that are tightly controlled by LexA will be

turned on. For example, the full induction of umuC and

umuD is not observed until 20 min after 40 J m�2 UV

irradiation (Courcelle et al., 2001). Similar to LexA, the

protein encoded by umuD also has a latent ability to auto-

digest, and the auto-digestion is strongly stimulated by the

interaction between the RecA/ssDNA nucleoprotein fila-

ment and UmuD (Nohmi et al., 1988). UmuC and a

posttranslationally processed form of UmuD (UmuD0) serve

as a mutagenic lesion-bypass DNA polymerase (PolV). This

last response allows the survival of E. coli after severe DNA

damage, but at the expense of introducing errors into the

genome.

In the SOS regulatory network, the transcript level of key

regulatory genes recA and lexA is also regulated by LexA,

thus forming a delicately controlled circuit (Brent &

Ptashne, 1980; Little et al., 1981; Brent, 1982). recA has one

SOS box positioned between the � 35 and � 10 regions of

the promoter, and a tandem pair of SOS boxes is located in

the lexA promoter region (Brent & Ptashne, 1981; Little

et al., 1981; Schnarr et al., 1991). LexA can bind to these SOS

boxes to prevent the initiation of transcription. Relative

binding affinity experiments reveal that LexA binds to the

recA operator more strongly than to the lexA operator and

many other operators of SOS regulon genes such as uvrA,

uvrB and uvrD (Brent & Ptashne, 1981; Peterson & Mount,

1987; Schnarr et al., 1991). Accordingly, it allows an inter-

mediate inducible state, bridging an uninduced state and a

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fully induced state (Little, 1983). A low amount of inducing

signal can thus lead to the activation of some of the SOS

functions, such as uvr1-dependent NER, without substan-

tial amplification of the RecA protein. When the inducing

signal continues to accumulate, recA will be induced, result-

ing in a full induction of SOS regulon genes. Meanwhile, due

to the repression of lexA by LexA itself, the SOS response

system is robust to prevent substantial induction caused by

very small amounts of inducing signal. Furthermore, the

strong repression of recA by LexA and the constant expres-

sion of LexA in SOS response ensure a fast return to the

uninduced state once the level of the inducing signal begins

to decrease (Walker, 1984).

In addition to the two central regulators RecA and LexA,

recent studies reveal that some other proteins are also

involved in the subtleties of SOS induction. All these

proteins appear to affect SOS regulation by modulating

RecA-ssDNA stability. RecX can block the assembly of the

RecA-ssDNA filament while not affecting the disassembly

through capping the assembly ends (Drees et al., 2004). As a

result, it strongly inhibits RecA-mediated DNA strand

exchange, ATPase and coprotease activities. The recX gene

is located 76-bp downstream of recA, and these two genes

belong to the same operon. Although recX is cotranscribed

with recA, recX transcription is downregulated with respect

to recA by an intrinsic transcription terminator that is

located between the recA and recX coding sequences. Despite

the presence of this terminator, a recA–recX message result-

ing from transcriptional read-through is detected at a level

of 5–10% of the recA message (Pages et al., 2003). RecX is

barely detected during vegetative growth, but robust expres-

sion of recX is observed after treating cells with DNA-

damaging agents (Stohl et al., 2003). The maximal recX

expression is observed at a later time than maximal expres-

sion of RecA after UV irradiation (Courcelle et al., 2001). All

these observations suggest that RecX is likely involved in the

subtle regulation that helps shut off the SOS response.

Early studies have reported that DinI can destabilize the

RecA-ssDNA filament when its concentration is 50–100-fold

above the natural RecA concentration, and it inhibits all

activities of RecA (Yasuda et al., 1998; Voloshin et al., 2001).

Therefore, DinI was initially thought to aid the return of

SOS-induced cells to a steady state. In contrast, recent

research has shown that DinI is also able to stabilize the

RecA-ssDNA filament to prevent its disassembly when

present at concentrations that are stoichiometric with or

somewhat greater than those of RecA (Lusetti et al., 2004).

Furthermore, DinI-mediated stabilization affects RecA-

mediated UmuD cleavage rather than RecA-mediated ATP

hydrolysis and LexA coprotease activities (Lusetti et al.,

2004), indicating that DinI could have a biological role in

fine-tuning the activity of UmuD in order to limit SOS

mutagenesis.

Role of DNA helicases and nucleases in SOSinduction

A critical step in the SOS response is the production of a

RecA-ssDNA filament, and this step requires DNA helicases

and nucleases. Either the RecFOR or the RecBCD pathway is

necessary for the SOS response after UV irradiation

(Ivancic-Bace et al., 2006). In E. coli, SOS induction im-

mediately after UV irradiation is dependent on the RecFOR

pathway. The RecFOR pathway includes the DNA helicase

RecQ, the nuclease RecJ and the RecFOR complex, which

facilitates RecA loading (Lusetti et al., 2006). It has been

suggested that the pathway may aid RecA binding to ssDNA

gaps. RecQ appears to be needed for fast degradation of the

LexA repressor (Hishida et al., 2004). This observation leads

to a model in which RecQ unwinds the template duplex in

front of a stalled fork on the leading strand, and then

switches over to the lagging strand to generate ssDNA on

the leading strand template, allowing formation of the RecA

filament in the 50–30 direction for SOS induction (Heyer,

2004; Hishida et al., 2004).

RecBCD, also known as Exonuclease V, contains helicase,

50–30 exonuclease, and RecA-loading activities. It can di-

rectly load RecA on the processed double-strand break

(DSB) ends (Singleton et al., 2004). SOS induction after

UV irradiation in recFOR mutants is not completely elimi-

nated but is delayed, and this induction is dependent on the

RecBCD enzyme (Thoms & Wackernagel, 1987; Hegde et al.,

1995; Whitby & Lloyd, 1995; Renzette et al., 2005). Accord-

ingly, it has been proposed that SOS induction requires

RecBCD when the DSB ends appear later because of NER

and replication fork collapse after UV irradiation (Ivancic-

Bace et al., 2006).

SOS-independent DNA damage induction inbacteria

Bacteria also possess damage-inducible systems that are

independent of the SOS response. For example, alkylating

or oxidative agents induce not only the SOS response but

also other more specific DNA damage responses.

Following exposure to a sublethal dose of an alkylating

agent (e.g. N-methyl-N0-nitro-N-nitrosoguanidine, MNNG),

bacteria such as E. coli manifest a pronounced resistance to

both lethal and mutagenic effects caused by a much higher

dose of the same or a similar alkylating agent (Jeggo et al.,

1977). The resistance is due to the induction of one or more

genes in response to low levels of the alkylating agents; the

protein products of these genes can directly remove the

alkylated adducts in DNA (Samson & Cairns, 1977). There-

fore, this regulatory pathway is called the adaptive response

to alkylation damage.

In E. coli, the Ada protein possesses dual functions and

plays a pivotal role in the adaptive response: it is a


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methyltransferase that removes alkyl groups from the

methylated base and a strong transcriptional activator of

several genes. The 39-kDa Ada protein is comprised of

two functional domains linked by a hinge region. The

C-terminal 19-kDa domain is capable of acquiring the

methyl group from a mutagenic O6-methylguanine or

O4-methylthymine to its Cys321 residue. The N-terminal

20-kDa domain specifically catalyzes the transfer of methyl

groups from methyl phosphotriester lesions to its Cys38

residue in a direct and irreversible way (Demple et al., 1985;

Nakabeppu & Sekiguchi, 1986; Teo et al., 1986; Lindahl

et al., 1988; Myers et al., 1993b; Takinowaki et al., 2006).

Methylation of the Cys38 triggers a conformational change

in the Ada protein, thereby dramatically (up to 1000-fold)

enhancing the sequence-specific DNA-binding affinity of

the Ada protein to the promoter regions of its own gene,

ada, and other alkylation resistance genes including alkA,

aidB and alkB (Teo et al., 1986; Sakumi & Sekiguchi, 1989;

Akimaru et al., 1990; Myers et al., 1993a; Sakashita et al.,

1993; Takinowaki et al., 2006). The transcriptional regula-

tory element (Ada box) in the ada gene to which methylated

Ada binds has been defined precisely, with the sequence 50-

AAAGCGCA-30. Methylated Ada also displays binding affi-

nity for the sequences 50-AAANNAAAGCGCA-30 and 50-

AAT(N)6GCAA-30 in the alkA and aidB promoter regions,

respectively (Teo et al., 1986; Nakamura et al., 1988; Landini

& Volkert, 1995).

Because the alkylation of Ada is irreversible, how is the

adaptive response switched off? Several mechanisms have

been proposed. One suggests that in the absence of alkyla-

tion adducts, the activated Ada is simply diluted by cell

divisions (Lindahl et al., 1988). Research has revealed that

physiologically relevant high concentrations of unmethy-

lated Ada are able to inhibit the activation of ada transcrip-

tion by methylated Ada, both in vitro and in vivo (Saget &

Walker, 1994). It is also possible that activated Ada is

proteolytically degraded. The proteolytic cleavage of acti-

vated Ada in the hinge region linking the N-terminal to

C-terminal domain of the protein might downregulate the

expression of ada. Consistent with this model, a methylated

20 kDa Ada can bind to the ada promoter; however, it does

not facilitate further binding of RNA polymerase to the

promoter nor does it promote ada transcription in vitro

(Akimaru et al., 1990).

The expression of genes responding to oxidative DNA

damage can be induced by the presence of reactive oxygen

species (ROS). Two regulatory responses induced by ROS

have been identified in E. coli – one controlled by soxRS and

the other by oxyR. In the soxRS regulatory system, SoxR

serves both as a sensor and as an activator. In the presence of

ROS, SoxR forms 2Fe–2S centers, which convert SoxR to an

active state. The activated SoxR induces transcription of

soxS, a positive regulator that stimulates transcription of

superoxide-responsive genes. Upon relief of oxidative stress,

SoxR is rapidly converted to its transcriptionally inactive

form, thus turning off the response (Wu & Weiss, 1992;

Hidalgo & Demple, 1994; Hidalgo et al., 1995; Ding et al.,

1996; Gaudu et al., 1997; Volkert & Landini, 2001). The oxyR

regulatory system is in response to hydrogen peroxide

(H2O2). H2O2 activates the transcriptional activity of OxyR

through formation of a disulfide bond between two of its

cysteine residues. Activated OxyR induces the transcription

of several oxidative stress genes (Zheng et al., 1998; Aslund

et al., 1999; Volkert & Landini, 2001).

Transcriptional responses to DNA damagein S. cerevisiae

From the above analyses, it appears that bacteria possess

transcriptional responses to specific types of DNA lesions as

well as general DNA damage, and that these regulatory

pathways are fairly well understood. In contrast, DNA

damage response in eukaryotes is apparently more complex

than in prokaryotes. Rather than direct cleavage of tran-

scription repressors, eukaryotes most likely use posttransla-

tional modifications, such as phosphorylation and

ubiquitination, and a sophisticated signal transduction

cascade to achieve gene regulation in response to DNA

damage. Here, we attempt to summarize the current knowl-

edge of mechanisms of DNA damage response in S. cerevi-

siae, a unicellular lower eukaryotic model organism.

Following a historic overview of identification and charac-

terization of genes whose transcript levels are altered follow-

ing treatment of cells with DNA-damaging agents, this

review focuses on regulatory responses that are either

common or unique to some well-studied genes.

Genome-wide DNA damage-induced expressionstudies in yeast

Following work in prokaryotes revealing that DNA damage

causes alterations in the transcriptional program of a cell, an

important initiative was to identify a comparable response

in the lower model eukaryote, S. cerevisiae. Although some

of these experiments were performed one gene at a time, a

precedent for systemic studies of transcription in bacteria

(Kenyon & Walker, 1980) and the emergence of high-

throughput technologies spurred the use of large-scale

screens for such studies in yeast. To originally identify some

of the genes that might be induced in response to DNA

damage in S. cerevisiae, two early studies utilizing different

genomic screens for DNA damage-induced genes are most

notable. The first was based on a transcriptional fusion

technique that is still widely used today for routine expres-

sion analysis. Using an experimental approach used pre-

viously in E. coli to screen for DNA damage-inducible genes

(Kenyon & Walker, 1980), Szostak and colleagues tested



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yeast genes as a random pool of lacZ fusions and screened

for coordinately regulated DNA damage-inducible genes

over various DNA damage treatments. From c. 8000 inde-

pendent clones, a total of six damage-inducible (DIN) genes

were identified, including one gene from a previous screen

(Ruby et al., 1983; Ruby & Szostak, 1985). Based on different

expression profiles of DIN genes to specific DNA-damaging

agents, the authors proposed at least two DIN classes. Such

grouping of inducible genes is analogous to the clustering of

large-scale data sets that is often necessary to manage

and interpret modern DNA microarray experiments. Mean-

while, McClanahan & McEntee (1984) used a differential

plaque hybridization strategy to identify genes whose tran-

script levels are altered after DNA damage treatment.

Radiolabeled cDNA probes were generated from poly(A)

mRNA harvested from control and treated cells, and

subsequently used for differential plaque hybridization.

By comparing the intensity of labeled probe bound between

the control and the treated samples, differential expression

of genes could be evaluated. Altogether, c. 9000

genomic clones were screened and four DNA damage-

responsive (DDR) genes were identified. The experimental

approach also enabled the identification of another set of

genes, with decreased expression in response to DNA

damage treatment. This latter finding testifies to the

complexity of the DNA damage response in eukaryotes,

because no such deactivation of genes was ever found in the

E. coli SOS response (Walker, 1984).

These two early screens were significant to the field of

transcriptional response to DNA damage in yeast. Not only

were they among the first to demonstrate altered levels of

transcripts in S. cerevisiae after DNA damage insult but also

they were carried out in a genome-wide manner and

foreshadowed some of the benefits and complications

associated with analogous DNA microarray experiments

that would follow. The powerful yet straightforward experi-

mental global approaches also paid dividends by consider-

ably expanding the number of genes known to be induced by

DNA damage at the time. Together, 10 new DNA damage-

inducible (DIN/DDR) and four DNA damage-repressive

genes were identified. These studies allowed estimation of

total DNA damage-responsive genes. Because the genomic

library of Ruby and Szostak likely contained only c. 500 yeast

genes representing about 8% of the genome, extrapolation

led to an estimation of roughly 80 DNA damage-inducible

genes in yeast. Significantly, the two independent screens

yielded nonoverlapping data sets, indicating that neither

screen was saturated and that the number of DNA damage-

inducible genes was probably underestimated. Thus, even in

the earliest stages of the studies of yeast DNA damage-

induced gene expression, it was obvious that S. cerevisiae

would have a much larger repertoire of DNA damage-

induced genes than E. coli.

With the exception of these studies, the identification

of DNA damage-inducible genes in the premicroarray era

was performed on a gene-by-gene basis, and the entire

complement of DNA damage-inducible genes grew very

slowly. A comprehensive list of these genes, including

relevant references, has been compiled and tabulated

(Friedberg et al., 1995) with references herein (McDonald

et al., 1997; Basrai et al., 1999; Bennett, 1999; Brusky

et al., 2000; Fry et al., 2003; Zaim et al., 2005; Fu & Xiao,

2006).

DNA microarray analyses in S. cerevisiae

Despite the above large-scale experimental approaches, a

plausible high-throughput and truly global approach was

not available until the emergence of genomic technologies.

In particular, DNA microarrays have provided the majority

of damage-induced gene expression data in budding yeast to

date. This wealth of expression data provided by DNA

microarrays has revealed a new realm for conceptualizing

regulatory networks in a global manner. In the following

paragraphs, we highlight only the first DNA microarray

studies of DNA damage-induced expression in yeast, and

note that there are numerous other significant studies that

cannot be feasibly covered in this review. Table 1 is provided

for an overview of genomic expression studies in S. cerevi-

siae focusing on DNA damage treatments or related gene

mutants. We refer the reader to the Yeast Functional

Genomics Database (http://yfgdb.princeton.edu/) for a na-

vigable compilation of these and related studies, 93 of which

are currently listed as genomic expression studies involving

stress.

The premiere microarray study on DNA damage-induced

expression in S. cerevisiae was published in 1999 (Jelinsky &

Samson, 1999). Representative sequences of 6218 yeast ORFs

were arrayed and cRNA probes were generated from untreated

cultures or cultures exposed to the DNA-alkylating agent

methyl methanesulfonate (MMS). As evidenced by the overlap

with the previous independent data mentioned above, the

results validated the use of DNA microarrays for damage-

induced expression analysis. Of the genes already known to be

induced by DNA-damaging agents by all other methods

combined, 86% (18 out of 21) were identified in the DNA

microarray experiment. The use of a single MMS dose and a

fourfold minimum cut-off likely explain the omission of the

three other previously identified inducible genes. Altogether, an

astonishing 325 yeast genes (c. 5% of entire yeast genes) were

found to be induced by MMS treatment, with a significant

portion (112) representing uncharacterized ORFs. In addition,

76 ORFs had reduced expression upon MMS exposure. The

reduced expression of some genes after DNA damage treatment

is reminiscent of the early DDR screen (McClanahan &

McEntee, 1984). Altogether, a single DNA microarray


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experiment augmented the complement of known DNA da-

mage-induced genes in S. cerevisiae by over 20-fold. Despite this

wealth of new data, however, meaningful conclusions regarding

transcriptional regulation were not possible with such an

abundance of raw, unorganized data.

A follow-up study (Jelinsky et al., 2000) provided a more

comprehensive analysis of transcriptional response to DNA

damage and demonstrated how large amounts of microarray

data can be manageably interpreted. This enabled the use of

a large dataset to address a specific hypothesis, namely that

MAG1 belongs to a novel regulatory network. The DNA

microarray data were computationally assembled into self-

organizing maps (SOMs) (Tamayo et al., 1999) that group

genes with similar expression profiles. Using SOMs, 18

different groups of genes were coregulated across various

experimental conditions. Of the genes in the SOM contain-

ing MAG1, it was noted that most contained an upstream

repressor sequence 2 element (URS2) previously identified

upstream of several genes involved in DNA repair and

metabolism (Xiao et al., 1993; Singh & Samson, 1995). It

was thus hypothesized and resolved that the genes within the

MAG1 SOM were regulated at the transcriptional level by

the same repressor element and constitute a regulatory

network. This study identified Rpn4 as a putative transcrip-

tional regulator of a large number of genes in response to

DNA damage. The fact that Rpn4 serves as a 26S protea-

some-associated protein as well as a transcriptional factor

that regulates at least 26 out of 32 proteasomal genes

(Mannhaupt et al., 1999) points to a possibility that

ubiquitin-mediated proteolysis may play a role in the

transcriptional response to DNA damage. Other studies

on the transcriptional response to DNA damage in yeast

using DNA microarray experiments have successfully ap-

plied similar methods of data management to test certain

hypotheses (see Table 1).

DNA damage checkpoint pathways andtranscriptional regulation

Both individual gene analyses and global transcriptional

studies in budding yeast indicate that many genes are

coordinately regulated in response to DNA damage.

Although transcriptional responses to specific damaging

agents result in distinct and signature profiles, the micro-

array data point to a general stress response pathway that

controls transcription. These observations are consistent

with previous reports based on individual gene analyses that

DNA damage and replication checkpoints are involved in a

transcriptional response to DNA damage.

The DNA damage checkpoint mutants were first isolated

by their failure to delay cell-cycle progression into mitosis

after irradiation with X-rays (Weinert & Hartwell, 1988;

Rowley et al., 1992; Weinert et al., 1994). These mutants

displayed increased radiation sensitivity, which could be

largely negated through reimposing an artificial arrest before

the M phase by an antitubulin agent. Hence, DNA damage

checkpoint was initially defined as a nonessential and

reversible response that slows down or arrests cell-cycle

progression in response to DNA damage, allowing time for

DNA repair (Hartwell & Weinert, 1989). At present, about

20 genes in budding yeast have been identified or antici-

pated to be involved in DNA damage checkpoints (Table 2)

(Elledge, 1996; Nyberg et al., 2002; Friedberg et al., 2006).

Table 1. Genomic DNA microarray studies of transcriptional response to

DNA-damaging agents and DNA repair-associated mutations in Sacchar-

omyces cerevisiae

References Cell type Mutant Treatment

Jelinsky & Samson (1999) Haploid MMS

Jelinsky et al. (2000) Haploid MMS, MNNG,

BCNU, t-BuOOH,

4-NQO, IR, MMC

Haploid rpn4 MMS

Gasch et al. (2000) Haploid H2O2

Gasch et al. (2001) Haploid MMS, IR, H2O2

Haploid dun1 MMS

Haploid mec1 MMS, IR

Haploid crt1 NA

De Sanctis et al. (2001) Haploid rad53,

rad6

IR

Oshiro et al. (2002) Haploid MMS, HU, UV,

Ostapenko & Solomon

(2003)

Haploid ctk1 HU

Green & Johnson (2004) Haploid tup1 NA

van Attikum et al. (2004) Haploid ino80,

arp8

MMS

Keller-Seitz et al. (2004) Diploid mec1 Aflatoxin B1

(N7-guanine

DNA adducts)

Mercier et al. (2005) All types IR

John et al. (2005) Haploid Cigarette smoke

extract

Benton et al. (2006) Haploid MMS, IR

Guo et al. (2006) Diploid Aflatoxin B1

Kelly et al. (2006) Haploid Azinomycin B

(interstrand

crosslinks and

alkylating agent)

Marques et al. (2006) Haploid H2O2

Shenton et al. (2006) Haploid H2O2

Workman et al. (2006) Haploid 30 TFs MMS

Kugou et al. (2007) Diploid mre11,

rad50

NA

Fu et al. (2008) Haploid rad6,

rad18

MMS

MMS, methyl methanesulfonate; MNNG, N-methyl-N0-nitro-N-nitroso-

guanidine; BCNU, 1,3-bis(2-chlorothyl)-1-nitrosourea; t-BuOOH, tert-

butyl hydroperoxide; 4-NQO, 4-nitroquinoline-n-oxide; IR, ionizing radia-

tions; MMC, mitomycin C; HU, hydroxyurea; UV, ultraviolet irradiation.

NA, not applicable.



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These genes are required for different phases of the cell cycle

and for different types of lesions. Because inactivation of

damage checkpoint genes affects the transcriptional re-

sponse to DNA damage as well as other cell fates, it has been

well accepted that checkpoint pathways play critical roles in

addition to cell-cycle arrest (Zhou & Elledge, 2000).

It is now clear that the checkpoint pathway comprises

a subroutine integrated into the larger DNA damage

response that regulates multifaceted responses (Zhou &

Elledge, 2000). Besides arresting cell-cycle progression, the

damage checkpoint pathway has been shown to promote

DNA repair (Mills et al., 1999), control telomere length

(Ritchie et al., 1999), activate transcription (Elledge, 1996)

and trigger apoptosis in metazoan cells (Roos & Kaina,

2006). In a broad sense, the damage checkpoint coordinately

regulates DNA damage responses including transcriptional

regulation.

Based on the data from previous research, a putative

model is proposed for the signal transduction pathways in

response to DNA damage, consisting of sensors, transducers

(including mediators) and effectors (Bachant & Elledge,

1998). The sensors are proteins that initially sense the

damaged DNA and initiate the signaling response. Transdu-

cers can be activated by the DNA damage signal passed

down from the sensor’s and then amplify and relay the signal

to the downstream effectors. The ultimate effectors are

defined as proteins that execute the cellular response. In the

case of transcriptional regulation, the effectors are likely to

be transcription factors that recognize cis-acting elements in

the promoter of DNA damage-inducible genes.

DNA damage sensors in S. cerevisiae

Because of the complexity of the source and type of DNA

damage, as well as the different cell-cycle stages at which

damage may occur, DNA damage sensors and their intrinsic

interactions have not been well established. Much of our

knowledge comes from studies with damage and replication

checkpoints.

The budding yeast proliferating cell nuclear antigen

(PCNA)-like 9-1-1 complex composed of Rad17, Ddc1 and

Mec3 is loaded on the DNA damage site with the assistance

of the clamp loader Rad24–Rfc complex, which leads to

activation of the main damage response kinase Mec1

(Kondo et al., 1999; Rouse & Jackson, 2002; Majka &

Burgers, 2003; Majka et al., 2006). Therefore, Rad24–Rfc

and the 9-1-1 complexes together have been speculated to be

sensors. Mec1 forms a stable complex with Lcd1/Ddc2, and

this complex itself has been anticipated to be a sensor due to

the ability of Ddc2 to bind to ends of single- and double-

stranded oligonucleotides in vitro and to recruit Mec1 to

double-strand break (DSB) sites in vivo in a manner

independent of the 9-1-1 complex (Kondo et al., 2001; Melo

et al., 2001; Rouse & Jackson, 2002; Niida & Nakanishi,

2006). Another sensor candidate is Rad9. Rad9 is required

for the activation of DNA damage checkpoint pathways in

budding yeast, is phosphorylated after DNA damage in a

Mec1- and Tel1-dependent manner and subsequently inter-

acts with the downstream kinase Rad53 (Naiki et al., 2004).

Rad9 displays the ability to associate with DSBs and controls

the DNA damage-specific induction of some repair,

Table 2. DNA damage sensors and signal transducers in Saccharomyces cerevisiae and their Schizosaccharomyces pombe and mammalian orthologs

Category Protein function S. cerevisiae S. pombe Mammals

Damage sensors RFC-like clamp loader Rad24 Rad17 RAD17

Clamp loader Rfc2-5 Rfc2-5 RFC2-5

PCNA-like clamp Ddc1 Rad9 RAD9

Rad17 Rad1 RAD1

Mec3 Hus1 HUS1

S-phase sensors Pole Pol2/Dun2 Cdc20 PoleDpb11 Cut5/Rad4 TopBP1

Drc1/Sld2 Drc1 Unknown

DNA helicase Sgs1 Rqh1 WRN, BLM, RTS

Sensor/transducer PI3K-like kinases Mec1 Rad3 ATR

ATR partner Lcd1/Ddc2 Rad26 ATRIP

Transducers PI3K-like kinases Tel1 Tel1 ATM

Damage mediator Rad9 Crb2 BRCA1, MDC1, 53BP1

S-phase mediator Mrc1 Mrc1 Claspin

Tof1 Swi1 Unknown

Csm3 Swi3 Unknown

Kinase Rad53 Cds1 CHK2

Kinase Chk1 Chk1 CHK1

Pds1 Cut2 PTTG

Effector kinase Dun1 Unknown Unknown


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replication and recombination genes (Aboussekhra et al.,

1996; Naiki et al., 2004). Furthermore, Rad9 is thought to

act in a pathway distinct from that of Rad24/9-1-1 for

damage checkpoint (Lydall & Weinert, 1995) and transcrip-

tional response (de la Torre-Ruiz et al., 1998), suggesting

that Rad9 may serve as an alternative DNA damage sensor.

The above candidate sensors primarily respond to DNA

damage caused by UV, ionizing radiations and endogenous

DSBs that induce G1/S and G2/M checkpoints. They also

respond to excessive telomere ssDNA in the cdc13-1 mutant

(Lydall & Weinert, 1995) and the accumulation of DSBs due

to a defective Cdc9 DNA ligase (Weinert & Hartwell, 1993),

both inducing the G2/M checkpoint.

The intra-S checkpoint is experimentally activated by

modest doses of alkylating agents, such as MMS, that cause

replication–blocking lesions. In addition, inhibiting replica-

tion by means other than DNA damage, such as treatment

with the ribonucleotide reductase (Rnr) inhibitor hydro-

xyurea, can activate similar cellular responses. Allele-specific

mutants in the catalytic subunit of DNA polymerase e (Pol2)

are defective in MMS- and hydroxyurea-induced RNR3

(Navas et al., 1995) and MAG1 (Zhu & Xiao, 1998) expres-

sion. Hence, Pol2 and its interacting partners Dpb11 and

Drc1 may serve as S-phase sensors. It is of great interest to

note that Dpb11 has been reported to interact with the Ddc1

subunit of 9-1-1 (Wang & Elledge, 2002), and that physcial

interactions between Pole and the human homologs of

Rad24/9-1-1 were also reported (Makiniemi et al., 2001;

Post et al., 2003), indicating that the damage signal may be

relayed between sensors. Recently, the helicase Sgs1, which is

a member of the E. coli RecQ helicase subfamily that includes

mammalian homologs of human BLM, WRN and RTS,

responsible for the Bloom, Werner and a subset of Roth-

mund–Thomson syndromes, respectively, was proposed as a

candidate sensor. Sgs1 possesses ATPase activity and prefer-

entially binds to branched DNA substrates (Bennett et al.,

1998, 1999). Sgs1 might be involved in the creation of the

signal for checkpoint activation, perhaps by resolving aber-

rantly paired double helices (Cobb et al., 2002). It functions

in the same epistasis group as Pole to activate Rad53 in the

presence of hydroxyurea, and this signaling pathway acts in

parallel to that of Rad17 and Rad24 (Navas et al., 1995; Frei

& Gasser, 2000). However, it remains unclear whether

putative S-phase-specific sensors like Pole and Sgs1 recog-

nize a subset of DNA damage or their activities lead to

common DNA damage intermediates such as ssDNA (like

RecA in the SOS response).

The above checkpoint proteins are placed in the sensor class

primarily based on genetic and biochemical properties or

inference, as little direct biochemical evidence yet exists to

support their sensor role. In principle, checkpoint sensors are

capable of directly recognizing DNA lesions or interacting with

pre-existing DNA damage repair proteins specific for indivi-

dual lesions. In this context, DNA repair proteins may serve as

damage sensors for both checkpoint and/or transcriptional

responses. A good example is the interaction between NER and

DNA damage checkpoint in response to UV treatment. A

genetic screen for mutants defective in the activation of

checkpoint following UV lesions but not other types of damage

resulted in the identification of RAD14, which is absolutely

required for the UV-induced damage checkpoint. Interestingly,

the checkpoint activation requires processing of UV lesions by

the NER complex instead of mere lesion recognition by Rad14,

and Rad14 physically interacts with Ddc1 (Giannattasio et al.,

2004). Similarly, the Mre11/Rad50/Nbs1 (MRN) DSB-binding

complex is required for the activation of ATM kinase in

mammalian cells (Lee & Paull, 2005), although a similar

function has not been reported for the corresponding Mre11/

Rad50/Xrs2 (MRX) complex in budding yeast. In both the

above cases, processing of UV-induced lesions by NER and

DSBs by MRN/MRX results in ssDNA regions that may serve

as an ultimate damage signal.

DNA damage transducers in S. cerevisiae

Considerable efforts have been made to understand signal

transducers in the damage and replication checkpoint path-

ways, and the same signal transduction cascade likely func-

tions in the transcriptional response to DNA damage. Most

of the transducers are protein kinases that, once activated by

the DNA damage signal passed down from the sensors,

amplify and relay the signal to the downstream effectors. In

S. cerevisiae, checkpoint protein kinases Mec1, Rad53 and

Dun1 are necessary for the transcriptional response of most,

if not all, genes to DNA damage, and they appear to be

central transducers in the regulation network (Zhou &

Elledge, 1993; Allen et al., 1994; Kiser & Weinert, 1996;

Gasch et al., 2001). The initiator of this signal transduction

kinase cascade in budding yeast appears to be Mec1 and its

regulatory subunit Lcd1/Ddc2, which is a serine/threonine

protein kinase belonging to the phosphatidylinositol-3-

kinase (PI3K) family. Mec1 is required for the activation of

Rad53 through phosphorylation of consensus PI3K sites

within Rad53 (Pellicioli & Foiani, 2005; Ma et al., 2006).

However, efficient and direct phosphorylation of Rad53

by Mec1 is only observed in the presence of Rad9 in vitro,

and the stimulatory activity of Rad9 requires both phos-

pho- and FHA-dependent interaction with Rad53, which

allows Rad53 to be recognized as a substrate for Mec1

(Sweeney et al., 2005). Hence, Rad9 is defined as a

checkpoint mediator (or adaptor) that serves to facilitate

and amplify signals. Rad9 functions primarily at G1/S and

G2/M, while Mrc1 and its mammalian homolog claspin

has been proposed to function as an S-phase or a replica-

tion checkpoint mediator. Mrc1 and Tof1 form a stable

replication-pausing complex at the stalled replication fork,



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which is required to anchor subsequent DNA repair events

(Katou et al., 2003).

Activation of Rad53 is an essential intermediary step in

yeast DNA damage responses that include delaying cell-cycle

progression, promoting repair processes, stabilizing stalled

replication forks and regulating transcription (Branzei &

Foiani, 2006). The kinase Dun1 is one of the identified

targets of Rad53, and the activation of Dun1 requires

phosphorylation by Rad53 (Chen et al., 2007). Dun1 was

originally identified as a DNA damage-uninducible (dun)

mutant defective in the transcriptional activation of genes

encoding Rnr in response to DNA damage (Zhou & Elledge,

1993), and was subsequently shown to be required for the

induction of other DNA damage-inducible genes (Basrai

et al., 1999; Zhu & Xiao, 2001; Fu & Xiao, 2006). Genomic

expression research showed that deletion of DUN1 affected

the expression of 4 1000 genes in response to MMS, and

the response in the dun1D mutant is largely the same as the

response seen in the mec1 mutant (Gasch et al., 2001),

suggesting that most of the Mec1-dependent effects on

genomic expression are mediated by the downstream Dun1

kinase. The mechanism used by Dun1 for DNA damage

induction appears to be rather diverse and requires further

investigation.

In response to DNA damage, the primary mechanism for

transcriptional regulation is likely to be at the stage of

transcription initiation. The downstream effectors are there-

fore expected to be transcription factors that directly influ-

ence transcription initiation. However, unlike damage

checkpoint transducers that appear to be aligned in a linear

cascade and control most if not all DNA damage-inducible

genes, the downstream effectors are clearly diverse and

dedicated to a particular gene or a group of genes (Fig. 1).

In the next section, selected examples are presented to

provide insights into the molecular mechanisms of DNA

damage-induced transcriptional response and their interac-

tion with the checkpoint cascade.

Mechanisms of transcriptional regulationof yeast genes in response to DNAdamage

RNR genes

Regulation of RNR genes is the best-known example of a

eukaryotic transcriptional response to DNA damage to date.

Rnr is an enzyme that converts nucleoside diphosphates

(NDPs) into deoxynucleoside diphosphates (dNDPs),

which represents the rate-limiting step in the production of

the four dNTPs for DNA synthesis and repair (Reichard,

1988; Elledge et al., 1993; Jordan & Reichard, 1998). Altered

levels or an imbalance of dNTP pools can lead to a higher

rate of spontaneous mutagenesis or cell death. Rnr is an

a2b2 tetramer, and four genes, RNR1-4, encode the subunits

of budding yeast Rnr: RNR1 and RNR3 encode a (large)

subunits, while RNR2 and RNR4 encode b (small) subunits.

Expression of all four genes is inducible at the transcrip-

tional level by a variety of DNA-damaging agents (Elledge

& Davis, 1987, 1989, 1990; Huang & Elledge, 1997), among

which RNR3 exhibits the highest level of induction, and

its transcriptional regulation has been examined most

extensively.

The transcriptional level of the RNR3 gene is very low

under normal conditions. However, when treating yeast cells

with DNA-damaging agents such as UV, MMS or 4-nitro-

quinoline-1-oxide (4-NQO), the transcript level of RNR3

can be increased 100–500-fold (Elledge & Davis, 1990). In

order to determine the mechanisms of induction, a series of

mutants have been isolated that cause constitutive expres-

sion of RNR3 (crt mutants) (Zhou & Elledge, 1992). These

negative regulators of RNR3 expression are divided into two

groups: indirect regulators that result in endogenous DNA

damage or a state of metabolic stress such as nucleotide

depletion that results in the upregulation of RNR3, and

Fig. 1. A working model of DNA damage-induced transcription in

budding yeast. In budding yeast, DNA damage-induced transcription is

controlled by signal transduction pathways including sensors, transdu-

cers and effectors. Mec1, Rad53 and Dun1 form the central kinase

cascade to regulate the DNA damage-induced transcription of most

genes examined. The phosphorylation of trans-acting factors (effectors)

may change their affinity for the corresponding cis-acting elements in

the promoter region of target genes, thus activating their transcription.

Only three sets of well-characterized genes are depicted. Solid ovals

indicate protein kinases. Dotted lines and question mark indicate that the

molecular mechanism of this step(s) remains unknown. Note that the

signal-sensor-transducer cascades are based on studies with cell-cycle

checkpoints and have not been extensively examined with transcriptional

regulation.


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direct regulators involved in the regulatory pathway, includ-

ing transcription factors. The CRT1, TUP1 (CRT4) and

SSN6 (CRT8) genes encode negative regulators that bind to

the RNR3 promoter. A second screen was carried out for the

mutations that disrupt the ability of DNA damage to induce

transcription of RNR3, and the corresponding genes were

designated DUN for DNA damage uninducible (Zhou &

Elledge, 1993). The nonessential serine/threonine protein

kinase Dun1 was isolated through this screen. Genetic

analysis of crt1, tup1 and ssn6 showed that these mutations

were epistatic to dun1, providing a strong genetic verifica-

tion that CRT1, TUP1 and SSN6 function downstream of

DUN1 (Huang et al., 1998). Combined with observations

that the Dun1 upstream kinases Mec1 and Rad53 are also

essential for the DNA damage-induced transcription of

RNR3 (Huang et al., 1998), the signal transduction pathway

for RNR3 becomes conceivable (Fig. 2).

It is now clear that in response to DNA damage or

replication blocks, the Rad53 protein kinase is activated in

a Mec1-dependent manner, and activated Rad53 further

phosphorylates the protein kinase Dun1. The Mec1-Rad53-

Dun1 kinase cascade culminates in the phosphorylation of

Crt1. Crt1 is a DNA-binding protein that recognizes a 13-bp

consensus sequence termed the X box. Multiple X box-

related sequences of different strength can be found in the

promoter region of all four RNR genes. Crt1 is able to

recruit the Tup1–Ssn6 general repressor complex to suppress

the transcription of RNR genes. In the presence of DNA

damage or replication blocks, Crt1 is phosphorylated in a

Dun1-dependent manner and this phosphorylation

abolishes its ability to bind X boxes, leading to the transcrip-

tional induction of RNR genes (Huang et al., 1998).

Surprisingly, multiple X boxes were also identified in the

CRT1 promoter. Data from mutated X boxes showed that

they confer CRT1-dependent repression on CRT1 itself

(Huang et al., 1998). The expression level of CRT1 is very

low under normal growth conditions but is inducible by

DNA-damaging agents. Interestingly, the X boxes in the

CRT1 promoter consist of one with a weak affinity for Crt1

and one with a strong affinity. Thus, it is speculated that a

weak induction of CRT1 occurs immediately following DNA

damage, which provides a buffer against spurious transcrip-

tional activation of the pathway. Delaying full activation

may ensure a rapid restoration of the basal repressed state

when the DNA damage is repaired (Huang et al., 1998),

which may serve as an autonomous regulatory circuit (Fig.

2). This mechanism is reminiscent of the autonomous

regulation of the LexA repressor in the E. coli SOS response.

Crt1 mediates repression by recruiting the general re-

pressor Tup1–Ssn6 complex to RNR promoters via its

DNA damage

Mec1

Rad53

Dun1

P

Pol

Crt10

X-box CRT1

Crt1

RNR3Ccr4

CRT1 mRNA

Crt1

Crt1

X-boxCrt1

P

P

P

P

RNR3X-box

Rpd

3

Wtm

2

Hos2

Stalled replication

Fig. 2. Summary of the transcriptional regulation of RNR3. In response to DNA damage or stalled replication, activation of the Mec1-Rad53-Dun1

kinase cascade results in phosphorylation of the repressor protein Crt1, and phosphorylated Crt1 loses its binding affinity for the RNR3 promoter. With

the assistance of Wtm1, Rpd3 and Hos2, the transcription of RNR3 is highly activated. The transcription of CRT1 is negatively regulated by itself and

positively regulated by Crt10. Ccr4 influences Crt1 protein abundance by controlling the CRT1 mRNA stability.



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N-terminus (Huang et al., 1998; Li & Reese, 2000). Tup1-

Ssn6 recruitment establishes a nucleosomal array over the

promoter of RNR3 with a positioned nucleosome occupying

the TATA box to block access by the general transcriptional

machinery (Li & Reese, 2000, 2001; Sharma et al., 2003). The

derepression of RNR3 correlates with the disruption of the

nucleosome position. In response to DNA damage signals,

hyperphosphorylated Crt1 loses the ability to bind to the

RNR3 promoter, and the Tup1–Ssn6 complex is not re-

cruited to the promoter region. Consequently, the chromatin

structure is remodeled and thus increases the accessibility of

DNA to transcription factors. The chromatin remodeling at

the RNR3 promoter requires a number of general transcrip-

tional factors, such as TBP-associated factors (TAFIIS), and

RNA polymerase II. Furthermore, the remodeling is also

dependent on the SWI/SNF complex, which possesses a

DNA-stimulated ATPase activity and can destabilize histo-

ne–DNA interactions in an ATP-dependent manner (Sharma

et al., 2003). The preinitiation complex components TFIID

and RNA polymerase II aid in recruitment and retention of

the SWI/SNF complex to the RNR3 promoter.

Recently, it was reported that the expression of CRT1 is

also controlled by the novel regulator Crt10 (Fu & Xiao,

2006) (Fig. 2). CRT10 was identified through screening of

the S. cerevisiae deletion strains for hydroxyurea resistance.

Epistatic analysis indicates that CRT10 belongs to the CRT1

pathway. Deletion of CRT10 does not affect the transcript

level of TUP1 or SSN6 regardless of hydroxyurea treatment,

but significantly reduces the basal level as well as hydroxyur-

ea-induced expression of CRT1; thus, CRT10 appears to be a

positive regulator of CRT1 transcription (Fu & Xiao, 2006).

Furthermore, the dun1 mutation is epistatic to crt10 with

respect to both hydroxyurea sensitivity and RNR gene

expression. Interestingly, the expression of CRT10 itself is

induced by DNA-damaging agents and this induction

requires DUN1 (Fu & Xiao, 2006). The increased Crt10

activity may be required to bring Rnr activity back to a

normal level through increasing the expression of repressor

Crt1 once the DNA damage is repaired. Like CRT1, the

induction of CRT10 itself depends on DUN1, suggesting

that Crt10 functions downstream of Dun1 and forms

another component of the autoregulatory circuit to regulate

the expression of RNR genes (Fu & Xiao, 2006).

CCR4 encodes a component of the major cytoplasmic

deadenylase, which is involved in mRNA poly(A) tail short-

ening (Tucker et al., 2001). Cells defective in CCR4 display

particular sensitivity to the Rnr inhibiter hydroxyurea

(Woolstencroft et al., 2006). The ccr4 dun1 double mutants

exhibit synergistic sensitivity to hydroxyurea, and simulta-

neous overexpression of RNR2, RNR3 and RNR4 partially

rescues the hydroxyurea hypersensitivity of a ccr4 dun1

strain, implying that CCR4 and DUN1 function in different

pathways to regulate the activity of Rnr. Deletion of CRT1

suppresses hydroxyurea sensitivity of the ccr4 mutant, and

overexpression of CRT1 hypersensitizes ccr4 to hydroxyurea.

These observations lead to the conclusion that Ccr4 regu-

lates CRT1 mRNA poly(A) tail length and may thus subtly

influence the Crt1 protein abundance (Woolstencroft et al.,

2006) (Fig. 2).

More regulatory factors were found to be involved in the

transcriptional regulation of RNR genes. Wtm1 and Wtm2

have been reported to modulate the expression of RNR3

(Tringe et al., 2006). Moderate overexpression of both genes

or high-level expression of WTM2 alone upregulates RNR3-

lacZ in the absence of DNA damage. In response to hydro-

xyurea and g-rays, the expression level of RNR3 attenuated

45% in wtm2D mutants, but not in wtm1 mutants. Wtm2

was found to associate directly with the RNR3 promoter,

and the association correlates with its ability to increase

constitutive RNR3 expression. It remains unknown how

Wtm2 increases RNR3 transcription, although some obser-

vations hint that Wtm2 might enhance RNR3 transcription

by participating in chromatin remodeling (Tringe et al.,

2006). Furthermore, Wtm1 associates with and anchors the

Rnr2/Rnr4 complex in the nucleus in untreated cells. In

response to DNA damage or replication inhibition, the

interaction between Wtm1 and Rnr2/Rnr4 is disrupted and

this complex is released from the nucleus to the cytoplasm

(Lee & Elledge, 2006; Zhang et al., 2006). A recent report

shows that two histone deacetylases (HDAC), Rpd3 and

Hos2, are required for the transcriptional activation of

RNR3 in response to DNA damage (Sharma et al., 2007)

(Fig. 2). Although a direct association between Hos2 and the

RNR3 promoter has not been observed, Rpd3 was found to

specifically bind to X boxes in the RNR3 promoter region in

a Tup1- or a Crt1-independent manner. The HDAC activity

of Rpd3 and Hos2 is essential for the activation of RNR3.

The observation that the recruitment of RNA polymerase II

is dramatically reduced in the rpd3 hos2 mutant indicates

that Rpd3 and Hos2 activate the transcription by regulating

the assembly of the preinitiation complex or inducing

multiple rounds of RNA polymerase recruitment (Sharma

et al., 2007).

PHR1

PHR1 encodes a photolyase that specifically and exclusively

repairs pyrimidine dimers, which are the most abundant

lesions found in DNA following UV irradiation (Sancar,

1985). Interestingly, the transcription of PHR1 is induced

by various DNA-damaging agents such as MMS, MNNG,

UV irradiation, 4-NQO or g-ray (Robinson et al., 1986;

Sebastian et al., 1990) despite the fact that Phr1 only reverses

UV-induced pyrimidine dimers but not lesions induced by

other DNA-damaging agents. Three transcriptional regula-

tory elements have been defined within the PHR1 promoter


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region: an upstream activation sequence (UAS), an up-

stream repression sequence (URS) and an upstream essen-

tial sequence (UES) (Sancar et al., 1995). A 22-bp

interrupted palindrome comprises UASPHRI, and it is re-

sponsible for 80–90% of basal and induced expression. It

alone can activate transcription of a CYC1 minimal promo-

ter but does not confer damage responsiveness (Sancar et al.,

1995). URSPHR1 is defined to a 39-bp region that includes a

22-bp palindrome. Deletions or specific mutations within

URSPHR1 increase basal-level expression but decrease the

induction ratio. It functions as a strong URS and confers a

low-level damage inducibility when placed in the context of

a heterologous gene (Sancar et al., 1995). The UESPHR1 is

required for efficient derepression when URSPHR1 is present.

Deletion of URSPHR1 also eliminates the requirement for

UESPHR1 for transcriptional activation (Sancar et al., 1995).

Three proteins have been identified that regulate the

expression of PHR1 by binding to the upstream regulatory

elements. Ume6 is a bifunctional transcriptional regulator

involved in several metabolic pathways (Strich et al., 1994).

It is a positive regulator of PHR1 transcription and binds

specifically to the UASPHR1 (Sweet et al., 1997). Multiple

copies of Ume6 enhance the expression of PHR1; however,

deletion of UME6 reduces the expression of PHR1 during

vegetative growth, but only at a distinct cell-cycle phase

(Sweet et al., 1997; Sancar, 2000). Rph1 and Gis1, which are

35% identical to each other at the amino acid sequence level,

are two DNA damage-responsive repressors of PHR1 tran-

scription (Jang et al., 1999; Sancar, 2000). Both Rph1 and

Gis1 contain two putative zinc fingers that are 4 90%

identical overall and identical in the DNA-binding loop,

and they regulate the transcriptional response of PHR1

through binding to URSPHR1. Deletion of both RPH1 and

GIS1 is required to fully derepress PHR1 in the absence of

damage, suggesting that they are functionally redundant. In

vitro footprinting and binding competition studies indicate

that the sequence AG4 (C4T) within the URSPHR1 is the

binding site for Rph1 and Gis1 (Jang et al., 1999).

Induction of PHR1 is controlled by the DNA damage

signal transduction pathway. The serine and threonine

residues of Rph1 can be phosphorylated, and the phosphor-

ylation of Rph1 is increased in response to DNA damage.

The DNA damage-induced Rph1 phosphorylation requires

DNA damage checkpoint proteins Rad9, Rad17, Mec1 and

Rad53, indicating that the phosphorylation of Rph1 is under

the control of the Mec1-Rad53 DNA damage checkpoint

pathway (Kim et al., 2002). On the other hand, deletion of

DUN1, TEL1 or CHK1 does not affect the phosphorylation

of Rph1, indicating that PHR1 is regulated by a potentially

novel damage checkpoint that is distinct from the Mec1-

Rad53-Dun1 protein kinase cascade. Based on the results of

a coimmunoprecipitation assay, Rad53 does not appear to

interact physically with Rph1, indicating the existence of a

yet unknown kinase(s) in the Mec1-Rad53-Rph1 pathway

(Kim et al., 2002).

MAG1 and DDI1

MAG1 encodes a 3-methyladenine DNA glycosylase that

initiates the base excision repair pathway by removing lethal

lesions such as 3-methyladenine (Chen et al., 1989). MAG1

is not only induced by DNA-alkylating agents such as MMS

but also by UV, 4-NQO or hydroxyurea (Chen et al., 1990;

Chen & Samson, 1991; Xiao et al., 1993). By analyzing the

DNA sequence immediately upstream of MAG1, another

DNA damage-inducible gene, named DDI1 for DNA Da-

mage Inducible, was identified (Liu & Xiao, 1997). Similar

to MAG1, DDI1 is also induced by MMS, UV, 4-NQO and

hydroxyurea. Furthermore, both genes require a similar

dosage for maximum induction, and the induction profile

is similar (Liu & Xiao, 1997).

MAG1 and DDI1 lie in a head-to-head configuration and

are transcribed divergently. The expression of both MAG1

and DDI1 is controlled by common as well as distinct UAS

and URS elements, possibly through antagonistic mechan-

isms (Liu & Xiao, 1997). A UASMAG1 and a URSMAG1 have

been identified in the promoter region of MAG1 (Xiao et al.,

1993). Interestingly, fine mapping of the UASMAG1 sequence

reveals that it is located within the DDI1 protein coding

region, and the electrophoretic mobility shift assay using

labeled UASMAG1 as a probe detected sequence-specific

binding protein(s) (Liu & Xiao, 1997), although the identity

of the UASMAG1-binding protein has not been reported. The

expression of DDI1 is negatively regulated by a URSDDI1 in

its promoter region (Liu & Xiao, 1997). The intergenic

region between MAG1 and DDI1 also contains a cis-acting

element that coregulates the expression of both genes. In this

shared promoter region, UASDM, which contains two 8-bp

tandem repeat sequences 50-GGTGGCGA-30, is required for

the bidirectional expression of MAG1 and DDI1; deletion or

point mutations of this tandem repeat result in a reduced

basal level expression and significant reduction of damage-

induced expression (Liu & Xiao, 1997). Furthermore,

UASDM alone is able to confer a DNA damage response

when fused to a CYC1 promoter (Liu & Xiao, 1997),

indicating that this cis-acting element independently inter-

acts with a transcriptional factor(s). With a yeast one-hybrid

screen using UASDM as a bait, a transcriptional activator

called Pdr3 was isolated (Zhu & Xiao, 2004). Pdr3 binds to

UASDM in vivo and in vitro, and deletion of PDR3 reduced

both the basal level and the DNA damage-induced expres-

sion of MAG1 and DDI1. In addition, deletion of PDR3 does

not further affect MAG1 and DDI1 expression if UASDM is

deleted, indicating that UASDM is indeed the target for Pdr3

activation (Zhu & Xiao, 2004). Another transcriptional

activator, Rpn4, was shown to be required for MAG1



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(Jelinsky et al., 2000) and DDI1 (Zhu & Xiao, 2004)

expression; however, Rpn4 does not appear to bind UASDM.

Moreover, deletion of RPN4 does not alter MAG1 and DDI1

expression in the pdr3 mutant cells, suggesting that RPN4

acts upstream of PDR3 (Zhu & Xiao, 2004). Deletion of

PDR3 and RPN4 has no effect on the basal level or the DNA

damage-induced expression of PHR1, RNR2 or RNR3.

Meanwhile, Crt1 and Tup1/Ssn6, the repressors of the RNR

genes, and Rph1 and Gis1, the repressors of PHR1, are not

involved in the control of MAG1 expression (Zhu & Xiao,

2001). Hence, it becomes apparent that all three sets of well-

studied yeast damage-inducible genes (RNR, PHR1 and

MAG1-DDI1) have distinct regulators and that the regula-

tory mechanisms are also different from each other.

The expression of MAG1 and DDI1 is also controlled by

the DNA damage checkpoint. Mutation of POL2, MEC1 or

DUN1 reduces the DNA damage response of MAG1 (Zhu &

Xiao, 1998, 2001), suggesting that MAG1 is regulated by the

POL2-MEC1-RAD53-DUN1 checkpoint pathway. In con-

trast, DDI1 remains inducible in sad1-1 (rad53), dun1 or

mec1 mutants, but its induction is diminished in pds1 sad1-1

(rad53) or pds1 dun1 double mutants (Zhu & Xiao, 2001).

PDS1 encodes an anaphase inhibitor and functions down-

stream of CHK1; CHK1-PDS1 and RAD53-DUN1 may form

two parallel branches in the DNA damage checkpoints (Fig.

1) (Gardner et al., 1999; Schollaert et al., 2004). This

suggests that the CHK1-PDS1 and MEC1-RAD53-DUN1

checkpoint pathways may function redundantly in the

control of DDI1 expression (Zhu & Xiao, 2001).

A putative eukaryotic SOS response in buddingyeast?

Although cell-cycle checkpoints have been regarded as a

eukaryotic SOS response, their similarity to the bacterial

SOS response is limited. In budding yeast, the Rad51 protein

is a sequence homolog of E. coli RecA and possesses ssDNA-

binding and ATPase activities (Shinohara et al., 1992).

However, the only Rad51 enzymatic activity reported to date

is ATP- and ssDNA-dependent recombinase, and it is in-

volved in homologous recombination and homology-

mediated DNA repair and damage tolerance pathways (Sung,

1994; Paques & Haber, 1999; Gangavarapu et al., 2007).

Although RAD51 itself is DNA damage inducible (Shinohara

et al., 1992), deletion of RAD51 does not affect the expression

of the DNA damage-inducible genes examined (Y. Fu &

W. Xiao, unpublished results). Nevertheless, it remains

possible that there is a functional homolog of RecA in yeast

with respect to a transcriptional response but it shares no

sequence homology and enzymatic activity with RecA.

In a broad sense, E. coli RecA controls three important

cellular responses to DNA damage, namely homologous

recombination via RecBCD and RecFOR, TLS by working

with PolIV and PolV and the SOS response, which collec-

tively provides a survival mechanism when cells encounter

replication blocks and/or contain excessive ssDNA. The

above functions are reminiscent of the Rad6–Rad18 com-

plex in budding yeast. Rad6–Rad18 forms a stable complex

that is essential for the DNA postreplication repair (PRR)

pathway (Broomfield et al., 2001). The Rad6–Rad18 com-

plex possesses ssDNA-binding and ATPase activities (Bailly

et al., 1997) and assumes most if not all RecA functions to

coordinate broad cellular responses to DNA damage. Firstly,

Rad6–Rad18 as an E2–E3 ubiquitination complex mono-

ubiquitinates PCNA to promote Polz- and PolZ-mediated

TLS (Hoege et al., 2002; Stelter & Ulrich, 2003). Secondly,

this mono-Ub-PCNA is required for PCNA poly-ubiquiti-

nation via a Lys63 chain linkage for an error-free mode of

DNA damage tolerance (Hoege et al., 2002) reminiscent of

the RecFOR activity in E. coli (Chow & Courcelle, 2004).

Finally, although Rad6–Rad18 does not have homologous

recombinase activity like RecA, it may compete with the

Ubc9–Siz1 complex that sumorylates PCNA at the same

K164 residue; sumorylated PCNA recruits the DNA helicase

Srs2, which inhibits the recombinase activity of the yeast

RecA homolog Rad51 (Papouli et al., 2005; Pfander et al.,

2005). Hence, it would be of great interest to investigate

whether the Rad6–Rad18 complex also plays a role in an

SOS response and, if so, how it interacts with the damage

checkpoints.

Conclusions

In summary, transcriptional response to DNA damage is an

important process for cell survival of microorganisms under

such life-threatening stress. In bacteria such as E. coli,

the DNA damage-induced transcriptional regulation is

centrally controlled by an SOS response. All SOS-inducible

genes contain SOS boxes in their operator regions, and

their expression is repressed by the interaction between the

transcriptional repressor LexA and SOS boxes under

noninduced conditions. The DNA damage induction is

achieved through enhanced self proteolysis of LexA

assisted by ssDNA-activated RecA. In addition, bacteria also

possess regulatory mechanisms in response to specific DNA

damage.

DNA damage-induced transcriptional response in eukar-

yotic microorganisms is apparently more complicated and

less understood. Nevertheless, several conclusions can be

drawn from data analyses as presented in this review. Firstly,

the idea that eukaryotic microorganisms possess an SOS

response structurally conserved from bacteria has been

effectively ruled out. Secondly, in budding yeast, a large

number of yeast genes (more than 5%) are induced after

DNA damage. Almost all DNA damage-induced genes

individually investigated to date respond to a broad range


920 Y. Fu et al.

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of DNA-damaging agents regardless of whether the gene

function provides a survival value to the particular damage

(Birrell et al., 2002). This observation supports the hypoth-

esis that there may exist a coordinated signal transduction

pathway to regulate transcriptional response to DNA da-

mage. Thirdly, DNA damage checkpoint pathways variably

affect transcriptional response to DNA damage in budding

yeast and have been regarded as a functional analog of the

bacterial SOS response. Fourthly, unlike the SOS-inducible

genes in bacteria, few DNA damage-inducible genes in

budding yeast share a common promoter sequence, suggest-

ing that the downstream effectors are diversified. This is

consistent with reports that all three well-characterized sets

of damage-inducible genes in budding yeast are controlled

by different transcriptional regulators and furthermore,

their modes of regulation are different: while RNRs and

PHR1 are induced by derepression, MAG1 and DDI1 are

induced by activation. Fifthly, the search for a novel

mechanism(s) in eukaryotic microorganisms that may bet-

ter resume the RecA-mediated SOS response is underway

and may result in exciting insights into the eukaryotic SOS

response. Finally, although transcriptional response to DNA

damage is a convenient and useful indicator for the gene

function, transcriptional levels do not necessarily reflect

cellular protein levels, and this appears to be the case for

DNA damage response in budding yeast (Lee et al., 2007).

Hence, other posttranscriptional regulatory mechanisms

have to be taken into account when examining a particular

gene function and its product activity.

Acknowledgements

The authors wish to thank the laboratory members for

helpful discussion and Michelle Hanna for proofreading

the manuscript. This work is supported by the Natural

Sciences and Engineering Research Council of Canada

discovery grant OGP0138338 to W.X. Y.F. is a recipient of

the Arthur Smyth Memorial Scholarship from College of

Medicine, University of Saskatchewan.

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